Proteins carry out the majority of signaling, metabolic, and regulatory tasks necessary for life. As a result, a quantitative description of the proteomic state of cells, tissues, and fluids is crucial for assessing the functionally relevant differences between diseased and unaffected tissues, between cells of different lineages or developmental states, and between cells executing different regulatory programs. Although powerful high-throughput techniques are available for determining the RNA content of a biological sample, the correlation between mRNA and protein levels is low (1).
The preferred method for proteomic characterization is currently mass spectrometry. Despite its many successes, mass spectrometry possesses limitations. One limitation is quantification. Because different proteins ionize with different efficiencies, it is difficult to compare relative amounts between two samples without isotopic labeling (2). In ‘shotgun’ strategies for analyzing complex samples, the uncertainties of peptide assignment further complicate quantification, especially for low abundance proteins (3). A second limitation of mass spectrometry is its dynamic range. For unbiased samples that have not undergone prefractionation or affinity purification, the dynamic range in analyte concentration is roughly 102-103, depending upon the instrument (4). This is problematic for complex samples such as blood, where two proteins whose levels are measured in clinical laboratories (albumin and interleukin-6) can differ in abundance by 1010 (5). Another limitation is the analysis of phosphopeptides, due to the loss of phosphate in some ionization modes. The power of proteomic approaches would increase dramatically with the introduction of a more quantitative high-throughput assay possessing greater dynamic range.
One promising technology for the analysis of proteins in a sensitive and quantitative manner was developed by Mitra et al (7). This technology, referred to as Digital Analysis of Proteins by End Sequencing or DAPES, features a method for single molecule protein analysis. To perform DAPES, a large number (ca. 109) of protein molecules are denatured and cleaved into peptides. These peptides are immobilized on a nanogel surface applied to the surface of a microscope slide and their amino acid sequences are determined in parallel using a method related to Edman degradation. Phenyl isothiocyanate (PITC) is added to the slide and reacts with the N-terminal amino acid of each peptide to form a stable phenylthiourea derivative. Next, the identity of the N-terminal amino acid derivative is determined by performing, for example, 20 rounds of antibody binding with antibodies specific for each PITC-derivatized N-terminal amino acid, detection, and stripping. The N-terminal amino acid is removed by raising the temperature or lowering pH, and the cycle is repeated to sequence 12-20 amino acids from each peptide on the slide. The absolute concentration of every protein in the original sample can then be calculated based on the number of different peptide sequences observed.
The phenyl isothiocyanate chemistry used in DAPES is the same used in Edman degradation and is efficient and robust (>99% efficiency). However, the cleavage of single amino acids requires strong anhydrous acid or alternatively, an aqueous buffer at elevated temperatures. Cycling between either of these harsh conditions is undesirable for multiple rounds of analysis on sensitive substrates used for single molecule protein detection (SMD). Thus, there is a need in the art for improved reagents and methods for the parallel analysis of peptides in single molecule protein detection (SMD) format.